SYNAPSE 69:148–165 (2015)

Female Protection From Slow-Pressor Effects of Angiotensin II Involves Prevention of ROS Production Independent of NMDA Receptor Trafficking in Hypothalamic Neurons Expressing Angiotensin 1A Receptors JOSE MARQUES-LOPES,1* MARY-KATHERINE LYNCH,1 TRACEY A. VAN KEMPEN,1 ELIZABETH M. WATERS,2 GANG WANG,1 COSTANTINO IADECOLA,1 VIRGINIA M. PICKEL,1† AND TERESA A. MILNER1,2†* 1 Brain and Mind Research Institute, Weill Cornell Medical College, New York, NY 2 Harold and Margaret Milliken Hatch Laboratory of Neuroendocrinology, Rockefeller University, New York, NY

KEY WORDS

paraventricular nucleus; sex differences; angiotensin; blood pressure

ABSTRACT Renin–angiotensin system overactivity, upregulation of postsynaptic NMDA receptor function, and increased reactive oxygen species (ROS) production in the hypothalamic paraventricular nucleus (PVN) are hallmarks of angiotensin II (AngII)-induced hypertension, which is far more common in young males than in young females. We hypothesize that the sex differences in hypertension are related to differential AngII-induced changes in postsynaptic trafficking of the essential NMDA receptor GluN1 subunit and ROS production in PVN cells expressing angiotensin Type 1a receptor (AT1aR). We tested this hypothesis using slow-pressor (14-day) infusion of AngII (600 ng/kg/min) in mice, which elicits hypertension in males but not in young females. Two-month-old male and female transgenic mice expressing enhanced green fluorescent protein (EGFP) in AT1aR-containing cells were used. In males, but not in females, AngII increased blood pressure and ROS production in AT1aR–EGFP PVN cells at baseline and following NMDA treatment. Electron microscopy showed that AngII increased cytoplasmic and total GluN1–silver-intensified immunogold (SIG) densities and induced a trend toward an increase in near plasmalemmal GluN1–SIG density in AT1aR–EGFP dendrites of males and females. Moreover, AngII decreased dendritic area and diameter in males, but increased dendritic area of small (1 mm) dendrites in females. Fluorescence microscopy revealed that AT1aR and estrogen receptor b do not colocalize, suggesting that if estrogen is involved, its effect is indirect. These data suggest that the sexual dimorphism in AngII-induced hypertension is associated with sex differences in ROS production in AT1aR-containing PVN cells but not with postsynaptic NMDA receptor trafficking. Synapse 69:148–165, 2015. VC 2015 Wiley Periodicals, Inc. INTRODUCTION The incidence of hypertension is greater in men than in young women; however, this condition is reversed at menopause (Martins et al., 2001; Wiinberg et al., 1995). In mice, administration of slowpressor doses of angiotensin II (AngII) induces hypertension in 2-month-old males, but not in age-matched females (Girouard et al., 2009; Marques-Lopes et al., 2014; Xue et al., 2013a,b). Increasing evidence points to sex differences in central cardiovascular control (Marques-Lopes et al., 2014; Pierce et al., 2009; Wang Ó 2015 WILEY PERIODICALS, INC.

Contract grant sponsor: NIH; Contract grant numbers: DA08259; AG016765; HL096571; AG059850; T32 DA007274.

HL098351;

*Correspondence to: Jose Marques Lopes, Robarts Research Institute, The University of Western Ontario, London, Ontario, Canada N6A 5K8. E-mail: [email protected] (or) Teresa A. Milner, Brain and Mind Research Institute, Weill Cornell Medical College, Room No. 307, 407 East 61st Street, New York, NY 10065, USA. [email protected] †

Virginia M. Pickel and Teresa A. Milner are co-senior authors.

Conflict of Interest: The authors have no conflict of interest. Received 7 August 2014; Accepted 23 December 2014 DOI: 10.1002/syn.21800 Published online 6 (wileyonlinelibrary.com).

January

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HYPOTHALAMIC SEX DIFFERENCES AND NMDA TRAFFICKING

et al., 2008; Xue et al., 2013a), and differential responses to hypertensive challenges in key brain areas could contribute to sex differences in the development and maintenance of hypertension. AngII plays a pivotal role in human hypertension through diverse mechanisms, and AngII administration is often used as a model of the disease (Biancardi et al., 2014; O’Callaghan et al., 2013; Paton et al., 2008). In the “slow-pressor” model of hypertension, circulating AngII modulates central cardiovascular control via blood–brain barrier-deprived circumventricular organs, for example, the subfornical organ (SFO; O’Callaghan et al., 2013). The SFO sends excitatory projections to the hypothalamic paraventricular nucleus (PVN), a key area in central modulation of blood pressure (Ferguson, 2009; Lind et al., 1982; Miselis, 1981). Excitatory SFO input to the PVN leads to local AngII release and increased expression of angiotensin Type 1 (AT1) receptor protein and mRNA (Sriramula et al., 2011; Wei et al., 2009; Wright et al., 1993). Likewise, renin–angiotensin system (RAS) overactivity in the PVN has been shown in spontaneously hypertensive rats (SHRs) and in AngII-induced hypertension (Sriramula et al., 2011; Veerasingham and Raizada, 2003; Wei et al., 2009). PVN infusion with AT1 or glutamate receptors blockers decreases blood pressure in hypertensive rats (Freeman and Brooks, 2007; Gabor and Leenen, 2012a, 2013). In accordance with an interaction between AngII and glutamate, AngII application in hypothalamic slices depolarizes PVN neurons through glutamate release from local interneurons (Latchford and Ferguson, 2004). Increased glutamatergic transmission in the PVN leads to the sympathoexcitation underlying the hypertension (Ferguson and Latchford, 2000; Gabor and Leenen, 2012b). The upregulation of postsynaptic NMDA receptor activity in PVN neurons enhances sympathetic outflow in SHRs and in AngII-induced hypertension (Li and Pan, 2010; Li et al., 2008, 2014; Wang et al., 2013; Ye et al., 2012). The augmented reactive oxygen species (ROS) production plays a crucial role in sympathoexcitation associated with AngIIdependent hypertension (Coleman et al., 2013; Jancovski et al., 2013; Wang et al., 2013). The central administration of NADPH oxidase inhibitors attenuates blood pressure elevation and ROS production in the PVN (Erdos et al., 2006). Prior work from our group has shown increased baseline and NMDA-evoked ROS production in PVN cells and enhanced association of GluN1 with NADPH oxidase in PVN dendrites of AngII-infused male mice (Wang et al., 2013). Sex differences in AngII-induced hypertension are associated with greater sympathoexcitation in males than in young females (Xue et al., 2005). We have recently shown that young female mice exhibit protection from AngII-induced hypertension associated with

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decreased NMDAR density in estrogen receptor (ER) b-containing PVN dendrites (Marques-Lopes et al., 2014). In the PVN, AT1 receptor and ERb are predominantly expressed in the parvicellular subregion in mouse and rat (Gonzalez et al., 2012; Hauser et al., 1998; Laflamme et al., 1998; Lenkei et al., 1997; Milner et al., 2010) and colocalize with corticotropinreleasing factor in rat (Aguilera et al., 1995; Laflamme et al., 1998). Thus, neurons containing AT1a receptor might colocalize ERb and be susceptible to local estrogen-mediated modulation of neuronal activity. Therefore, the current study sought to determine whether AngII infusion leads to differential changes in baseline and NMDA-evoked ROS production and in postsynaptic GluN1 density and trafficking in AT1a receptor (AT1aR)-expressing cells of the PVN in male and female mice. In addition, immunohistochemical analysis was performed to assess AT1 receptor and ERb colocalization in the PVN. Finally, retrograde tracing was used to evaluate if mouse PVN AT1aRcontaining neurons project to the median eminence (Hashimoto et al., 2004) and/or to the presympathetic neurons of the intermediolateral (IML) cell column of the spinal cord (Nunn et al., 2011). MATERIALS AND METHODS Animals General Experimental procedures were approved by the Institutional Animal Care and Use Committees of Weill Cornell Medical College and were in accordance with the 2011 Eighth Edition of the National Institute of Health Guide for the Care and Use of Laboratory Animals. The studies were conducted in Agtr1a bacterial artificial chromosome (BAC) transgenic mice that express AT1aR identified by enhanced green fluorescent protein (AT1aR–EGFP; Gonzalez et al., 2012). The details on the characterization of this mouse have been described previously (Gonzalez et al., 2012). Briefly, AT1aR–EGFP mice were originally developed by the GENSAT project (www.gensat. org) at the Rockefeller University (Gong et al., 2003). Hemizygote BAC-based AT1aR transgenic mice were originally on a FVB/N background and were backcrossed with C57BL/6 mice for six generations. Males and females between 2 and 3 months old at the beginning of the experiment were used. Mice were housed three to four animals per cage with 12:12 light/dark cycles with ad libitum access to food and water. All survival surgeries were done using isofluorane anesthesia (induction 5%; maintenance 1.5– 2% in oxygen). Estrous cycle determination Estrous cycle stage was determined using vaginal smear cytology (Turner and Bagnara, 1971) daily Synapse

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between 9:00 and 10:00 AM. In young (premenopausal) mice, estrous cycles last 4–5 days and consist of three primary phases: proestrus (high estrogen levels; 0.5–1 day), estrus (declining estrogen levels; 2– 2.5 days), and diestrus (low estrogen and progesterone levels; 2–2.5 days). Previous studies have shown variability in AT1 receptor density across the estrous cycle in the pituitary (Seltzer et al., 1992) and in the dorsomedial arcuate nucleus (Seltzer et al., 1993). Therefore, estrous cycle determination was performed to ensure that only females with two regular estrous cycles prior to beginning the experiment were used. No females in proestrus at Day 14 after minipumps implantation were included in the analyses. To control the effects of handling, male mice were removed from their cage and handled daily. Retrograde labeling of spinally projecting PVN neurons Male mice (N 5 3) were anesthetized as above, and their spinal cords were exposed at the T2–T4 level through dorsal laminectomy. Using a custom-made Hamilton syringe (Model 75 SN SYR, 5 ml, 32 gauge; Hamilton Company, Reno, NV), 1 ml of 4% Fluorogold (FG; Fluorochrome, Denver, CO) was pressureinjected into the IML region of the spinal cord, and the incision was sutured after the injection (Li et al., 2008; Marques-Lopes et al., 2014; Wang et al., 2013). Mice were euthanized 9 days after surgeries. Injection site was verified in spinal cord sections encompassing the T2–T4 region. Successful injections were centered in the IML. Limited lateral diffusion of FG into the intermediomedial nucleus was observed.

stress, the animals were handled by the same experimenter and at the same time of day throughout the study. Mice were euthanized at 1 day after the final SBP measurements (Coleman et al., 2013; MarquesLopes et al., 2014; Wang et al., 2013). ROS detection ROS production was determined using dihydroethidium (DHE). Superoxide oxidizes the cell-permeant DHE to 2-hydroxyethidium and other oxidation products (Zhao et al., 2003, 2005), which interact with DNA and are detectable by fluorescence microscopy. ROS production was measured in dissociated PVN cells from male and female mice after 14 days of vehicle or AngII infusion (n 5 13–16 cells per group, from five to seven mice each). Dissociation of PVN cells was performed with 90-min incubation as described previously (Coleman et al., 2013) with 0.02% pronase and thermolysin (Sigma-Aldrich, St. Louis, MO) in Mg21-free lactic acid-artificial cerebrospinal fluid. The identification of cells and ROS detection were performed as described previously (Girouard et al., 2009; Wang et al., 2013). Bath application of 100 mM NMDA was performed after a stable baseline measurement was achieved. NMDA is known to induce NOX2-dependent free radical production in the brain (Girouard et al., 2009). The increase in ROS signal induced by NMDA was expressed as the ratio of Ft/ Fo, where Ft is fluorescence after application of NMDA, and Fo is the baseline fluorescence in the same cell (Girouard et al., 2009; Wang et al., 2006, 2013). Immunocytochemical procedures

Labeling of neuroendocrine PVN neurons

Antisera

Male mice (N 5 3) were anesthetized as above and injected with 50 ll of 1% FG (Fluorochrome) into the tail vein using a 27G 1=2 insulin syringe (BD Biosciences, San Diego, CA), as described previously (Biag et al., 2012). Mice were euthanized 7 days after injections.

For labeling of EGFP, a chicken polyclonal antiGFP antibody (GFP-1020; Aves Labs, San Diego, CA) was used. The GFP antibody was generated against recombinant GFP and recognizes the gene product of EGFP-expressing transgenic mice (Encinas et al., 2006). The specificity of this antibody has been demonstrated by immunohistochemistry and Western blotting using GFP-expressing transgenic mice, resulting in one major band at 27 kDa (see data sheet for EGFP-1020 at www.aveslab.com). Moreover, the absence of labeling has been shown in brain sections from mice that lack EGFP (Milner et al., 2011; Volkmann et al., 2010). Previously, we showed that the distribution of AT1aR–EGFP cells in the brain closely corresponds to that obtained with labeling of AngII and AT1 receptor protein and mRNA (Gonzalez et al., 2012). For labeling of GluN1, a monoclonal mouse antiGluN1 antibody (clone 54.1; BD Biosciences) was used. Western blot analysis of a lysate from rat cortex probed with the anti-GluN1 antibody resulted in one

Slow-pressor AngII administration Mice were anesthetized as above, and osmotic minipumps (Alzet, Durect Corporation, Cupertino, CA) containing vehicle [saline 1 0.01% bovine serum albumin (BSA)] or AngII (600 ng/kg/min) were implanted subcutaneously in males and females (N 5 8–10 mice per group). Systolic blood pressure (SBP) was measured in awake mice by tail-cuff plethysmography (Model MC4000; Hatteras Instruments, Cary, NC), as described previously (Coleman et al., 2010), prior to (baseline) and 2, 5, 9, and 13 days after minipump implantation. The limitations of using tail-cuff plethysmography to measure SBP have been discussed previously (Marques-Lopes et al., 2014). To minimize Synapse

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major band at 120 kDa. HEK 293 cells transfected with cDNA-encoding GluN1 displayed similar results, whereas nontransfected cells resulted in no bands (see data sheet for anti-NMDAR1 at www.biosciences. com). The injection of rAAV-cre into the nucleus accumbens of floxed GluN1 mice leads to a 40% reduction of GluN1 immunoreactivity (Beckerman and Glass, 2012; Glass et al., 2013). To label ERb, an anti-ERb antibody raised in rabbits (Z8P; Zymed Laboratories, San Francisco, CA) was used. This antibody recognizes a peptide sequence in the C-terminus (amino acids 468–485) of the mouse ERb protein (Shughrue and Merchenthaler, 2001). The specificity for ERb has been shown by Western blotting, double labeled with mRNA using in situ hybridization, preadsorption control, and the absence of labeling in fixed brain sections prepared from ERb knock-out mice (Creutz and Kritzer, 2002; Shughrue and Merchenthaler, 2001). To label FG, an anti-FG antibody raised in guinea pig (Protos Biotech Corp, New York, NY) was used. We and others have shown selective staining of retrograde-labeled neurons using this antiserum in mouse PVN (Marques-Lopes et al., 2014), mouse hippocampus (Jinno and Kosaka, 2002), rat PVN (Perello and Raingo, 2013), and rat spinal cord (Polgar et al., 2007). Immunostaining is completely blocked by preincubation with FG or Fast Blue (see data sheet for NM-101 at www.protosantibody.com). For the labeling of arginine–vasopressin (AVP), a polyclonal anti-AVP antibody raised in guinea pig (Peninsula Laboratories, San Carlos, CA) was used. Immunostaining is completely abolished by preadsorption (Hundahl et al., 2010). Moreover, no staining is observed in the Brattleboro rat, which is unable to produce AVP due to a natural genetic mutation (Drouyer et al., 2010). Tissue preparation Fluorescence microscopy Spinally and tail vein-injected mice (described above) and additional female AT1aR–EGFP mice (N 5 5) were used to study if AT1aR and ERb colocalize in the PVN. Mice were deeply anesthetized with sodium pentobarbital (150 mg/kg, i.p.), and their brains were fixed by aortic arch perfusion sequentially with 2–3 ml saline (0.9%) containing 2% heparin followed by 30 ml of 4% paraformaldehyde (PFA) in 0.1 M sodium phosphate buffer (PB, pH 7.4). After the perfusion, the brains were postfixed for 24 h in 4% PFA at 4 C. Immunoelectron microscopy Three animals per group (males/females infused with AngII/saline) were deeply anesthetized with sodium pentobarbital as above and fixed by aortic arch perfusion sequentially with 2–3 ml saline (0.9%) containing 2% heparin followed by 30 ml of 3.75% acrolein and 2% PFA

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in PB (Milner et al., 2011). After the perfusion, brains were removed and postfixed for 30 min in 2% acrolein and 2% PFA in PB at room temperature. Brains were then cut into 5-mm coronal blocks using a brain mold (Activational Systems, Warren, MI) and sectioned (40 mm thick) on a VT1000X Vibratome (Leica Microsystems, Buffalo Grove, IL). Brain sections were stored at 220 C in cryoprotectant until immunohistochemical processing (Milner et al., 2011). To ensure identical labeling conditions between experimental groups for quantitative studies (Pierce et al., 1999), two sections per animal encompassing the region of the PVN [0.70–0.94 mm caudal to bregma; Fig. 1A (Hof et al., 2000)] were marked with identifying punches, pooled into single containers and then processed through all immunohistochemical procedures together. Dual-labeling immunofluorescence One in every three PVN sections was removed from the cryoprotectant, rinsed thoroughly in PB, and incubated in 1% BSA in PB for 2 h to minimize nonspecific binding of the antisera. The sections were incubated with combinations of appropriate primary antisera to (1) GFP (1:10,000) and ERb (1:1000), (2) EGFP and FG (1:2000), and (3) GFP, AVP (1:1200), and FG. Primary antisera were incubated in PB with 0.5% BSA and 0.1% Triton X-100. (1) The incubation in anti-ERb antibody was done at room temperature. At 24 h, the tissue was moved to 4 C. Anti-GFP antiserum was added to the primary antibody diluent at 96-h time point, and the tissue was incubated for additional 24 h. (2) The incubation in anti-GFP and anti-FG antisera was done at room temperature for 24 h. (3) The incubation in anti-GFP, anti-AVP, and anti-FG antisera was done at room temperature. At 24 h, the sections were moved to 4 C for additional 24 h. The sections were rinsed in PB and incubated for 2 h with combinations of appropriate secondary antibodies conjugated with Alexa Fluor dyes (1:400 dilution; Invitrogen-Molecular Probes, Carlsbad, CA). Alexa Fluor 488 goat anti-chicken IgG and Alexa Fluor 647 goat anti-rabbit pig IgG sections were mounted on gelatin-coated slides, air-dried, and coverslipped with SlowFade Gold reagent (InvitrogenMolecular Probes, Grand Island, NY). Dual-labeling electron microscopic immunocytochemistry Tissue sections were processed using a preembedding dual-immunolabeling protocol, as described previously (Milner et al., 2011). The tissue was treated with 1% sodium borohydride in PB for 30 min to neutralize reactive aldehydes and rinsed thoroughly in PB. Free-floating sections were immersed in 0.5% BSA in 0.1 M Tris saline (TS, pH 7.6) for 30 Synapse

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Fig. 1. Angiotensin Type 1a (AT1aR)–enhanced green fluorescent protein (EGFP)-labeled cells in the hypothalamic paraventricular nucleus (PVN). A: Diagram of a coronal section of the mouse brain showing a representative rostrocaudal level of the PVN containing AT1aR–EGFP (modified from Hof et al., 2000). B: Dense cluster of AT1aR–EGFP-containing cells in the PVN. Higher magnification (inset) shows that dendritic processes (arrow) of PVN neurons can be identified. C: Representative immunoelectron microscopic picture showing GluN1–silver-intensified immunogold (SIG) particles in an AT1aR–EGFP-labeled (immunoperoxidase) PVN dendrite. Immuno-

peroxidase labeling for AT1aR–EGFP (black arrow) was found throughout the cytoplasm of dendritic profiles. GluN1–SIG particles (black dots) were localized in the plasmalemma (thin magenta arrow), near the plasmalemma (magenta arrowhead), and in the cytoplasm (thick magenta arrow). Abbreviations: 3V, third ventricle; AHN, anterior hypothalamic nucleus; CEAm, central nucleus of the amygdala, medial part; CTX, cortex; SFO, subfornical organ; VMH, ventromedial nucleus of the hypothalamus. Scale bars: (B) and (C) 5 500 nm; inset, 25 lm.

min. The tissue was incubated at room temperature in a solution of anti-GluN1 (1:50) antiserum in TS with 0.01% BSA for 48 h. Anti-GFP (1:2500) antiserum was added to the primary antibody diluent at 24 h, and the tissue was moved to 4 C. For immunoperoxidase detection of GFP, the sections were placed for 30 min in goat anti-chicken IgG (1:400; Jackson ImmunoResearch, West Grove, PA), followed by 30-min incubation in avidin–biotin complex (Vector Laboratories, Burlingame, CA). After rinsing in TS, the bound peroxidase was visualized by reaction of the sections for 6–7 min in 3,30 -diaminobenzidine (DAB; Sigma-Aldrich Chemical, Milwaukee, MI) and hydrogen peroxide. For immunogold detection of GluN1, the DABreacted sections were rinsed and placed overnight in a 1:50 dilution of donkey anti-goat IgG with bound 1nm colloidal gold [Electron Microscopy Sciences (EMS), Fort Washington, PA]. The gold particles were fixed to the tissue in 2% glutaraldehyde in 0.01 M phosphate-buffered saline (PBS, pH 7.4) and rinsed in PBS followed by 0.2 M sodium citrate buffer (pH 7.4). The bound silver–gold particles were enhanced using a Silver IntenSE M kit (RPN491; GE Healthcare, Waukeska, WI) for 7 min. Tissue sections were postfixed in 2% osmium tetroxide for 1 h, dehydrated through a series of graded ethanols and propylene oxide, and flatembedded in Embed-812 (EMS) between two sheets of Aclar plastic. Ultrathin sections (70 nm thickness) from the PVN were cut with a diamond knife (EMS) using a Leica EM UC6 ultratome. The sections were collected on 400-mesh thin-bar copper grids (EMS)

and counterstained with uranyl acetate and lead citrate.

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Light microscopic analysis The sections containing the PVN (Fig. 1A) labeled for AT1aR–EGFP using immunoperoxidase were photographed with a Nikon 80i light microscope equipped with a micropublisher digital camera (Qimaging, Barnaby, British Columbia, Canada). Immunofluorescence microscopic analysis The photographs of spinal tract-tracing, ERb 1 AT1aR–EGFP labeling, and neuroendocrine PVN neurons were acquired using a Leica (Nussloch, Germany) confocal microscope. Z-stack analysis was used to verify the dually labeled neurons. The sections from spinal tract-tracing and ERb 1 AT1aR– EGFP reactions were collected from three different rostrocaudal levels encompassing the PVN [approximately 21.1, 21.6, and 22.1 mm from Bregma (Hof et al., 2000)]. Ultrastructural data analyses The sections were examined using a Tecnai transmission electron microscope. The images were collected at a magnification of 13,500. The profiles containing GluN1 with GFP immunoreactivity were classified as neuronal (soma, dendrites, axons, and terminals) or glial based on the criteria described by Peters et al. (1991). The dendritic profiles contained regular microtubular arrays and were usually postsynaptic to axon terminal profiles.

HYPOTHALAMIC SEX DIFFERENCES AND NMDA TRAFFICKING

An equal amount of tissue from each treatment group (9596 lm2 per group) was sampled for electron microscopic analysis. Immunoperoxidase labeling for GFP was distinguished as an electron-dense reaction product precipitate. Silver-intensified immunogold (SIG) labeling for GluN1 appeared as black electrondense particles. The criteria for field selection and the measures to avoid false-negative labeling of smaller profiles, variability between animals in each experimental group, and differential reagent sensitivity comparing SIG and immunoperoxidase labeling were performed as described before (Milner et al., 2011; Pierce et al., 1999). Tissue collection from each block was terminated when 50 images of dual-labeled dendritic profiles were taken. The analysis of GluN1– SIG density in non-AT1aR-containing dendrites was not performed, as these dendrites arise from a mixed population of neurons. The subcellular distribution and density of GluN1– SIG particles in AT1aR–EGFP-labeled dendrites was determined as previously described (Coleman et al., 2013). For this, the GluN1–SIG particle localization was categorized as (1) plasmalemmal, (2) near plasmalemmal (particles not touching but within 70 nm from the plasma membrane), or (3) cytoplasmic. The investigator performing the quantification of SIG particles was blinded to sex and experimental condition. Microcomputer Imaging Device software (Imaging Research, ON, Canada) was used to determine the cross-sectional diameter, perimeter, surface area, form factor, and major and minor axis lengths of each immunolabeled dendrite. The dendrites with an oblong or irregular shape (form factor value < 0.5) were excluded from the dataset. The parameters used for statistical comparisons were as follows: (1) number of plasmalemmal GluN1–SIG particles on a dendrite/dendritic perimeter, (2) number of near plasmalemmal GluN1–SIG particles, (3) number of cytoplasmic GluN1–SIG particles/dendritic crosssectional area, and (4) total GluN1–SIG particles (sum of plasmalemmal, near plasmalemmal, and cytoplasmic) in a dendritic profile. Dendrites were further divided into small (>1.0 lm) and large (